C57BL6/J mice (12–18 days old) were slightly anesthetized with isoflurane, decapitated, and the brain quickly removed and immersed in ice-cold high-sucrose “cutting solution” containing (in mM): 189.0 sucrose, 10.0 glucose, 26.0 NaHCO₃, 3.0 KCl, 5.0 Mg₂SO₄, 0.1 CaCl₂, 1.25 NaH₂PO₄·2H₂O. Coronal slices (350 μm thick) were cut with a Vibratome (Leica VT 1200S), then slices were incubated (>1 h, at room temperature, 25–27°C) in artificial cerebrospinal fluid (ACSF) which containing (in mM): 124.00 NaCl, 2.69 KCl, 1.25 KH₂PO₄, 2.00 Mg₂SO₄, 26.00 NaHCO₃, 2.00 CaCl₂, 10.00 glucose, 0.4 ascorbic acid and 0.3 sodium pyruvate. The pH was stabilized at 7.4 by bubbling the ACSF with carbogen (95% O₂, 5% CO₂). Slices were transferred to a 2 mL chamber fixed to an upright microscope stage (BX51WI; Olympus, Tokyo, Japan) equipped with infrared differential interference contrast video (DIC) microscopy and a 40X water-immersion objective and superfused at room temperature with carbogen-bubbled ACSF (2 mL/min). Cells were visualized under an Olympus BX50WI microscope. Patch-clamp recordings from the soma of L2/3 PNs of the barrel cortex were performed in the whole-cell voltage-clamp and current-clamp configurations. Patch pipettes were made from 1.50 OD/0.86 ID borosilicate glass capillaries (GC150F/10; Harvard Apparatus) using a P97 micropipette puller (Sutter Instruments) and had resistances of 4–8 MΩ when filled with an internal solution that contained (in mM): 120 K-Gluconate, 10 KCl, 10 HEPES, 0.5 EGTA, 4 Na₂-ATP, and 0.3 Na₃-GTP, 10 NaCl buffered to pH 7.2–7.3 with KOH. (280 mOsm) Recordings were performed in the current- or voltage-clamp modes using a Cornerstone PC-ONE amplifier (DAGAN, Minneapolis, MN). Pipettes were placed with a mechanical micromanipulator (Narishige, Tokyo, Japan). The holding potential was adjusted to −60 mV, and the series resistance was compensated to ~70%. L2/3 PNs located over the barrels (layer 4) were accepted only when the seal resistance was >1 GΩ and the series resistance (10–20 MΩ) did not change (>10%) during the experiment. Data were low-pass filtered at 3.0 kHz and sampled at 10.0 kHz, through an Axon Digidata 1440A interface board (Molecular Devices, Sunnyvale, CA). The pClamp software (Molecular Devices) was used to acquire the data. Chemicals were purchased from Sigma-Aldrich and Tocris Bioscience.

Synaptic responses were evoked with Pt/Ir concentric bipolar (OP 200 μm, IP 50 μm, FHC) stimulating electrodes (0.1 ms and 20–100 μA) connected by 2 silver-chloride wires to a Grass S88 stimulator and stimulus isolation unit (Quincy, United States). The stimulating electrode was placed at layer 4 of the barrel cortex. Single pulses (100 μs duration and 20–100 μA) were continuously delivered at 0.33 Hz.

In some experiments, after applying IGF-I as described above, spike-timing dependent plasticity (STDP) was induced (Figure 1). Induction of STDP was achieved by pairing pre- and postsynaptic action potentials 10 ms away. Presynaptic action potentials were evoked by electrical stimulation of basal afferent inputs. Stimulus intensity was adjusted to obtain 3–5 mV PSPs responses. Postsynaptic action potentials were elicited by a brief current injection through the recording pipette (5 ms, 200–400 pA). Basal PSPs were recorded for 10 min before the pre-post associations were induced (30, 20, 50 and 100 pairings were studied). After that, PSPs were recorded for another 50–60 min. Amplitude of PSPs 5 min before (−5 to 0 min interval) and 50 min after (45 to 50 min interval) the LTP_H induction protocol were measured to compare the extent of the PSP potentiation.

In all experiments, basal postsynaptic currents (PSCs) were recorded for 5 min at 0.33 Hz (voltage-clamp configuration), then basal postsynaptic potentials (PSPs) were recorded at 0.2 Hz (current-clamp configuration). After, the stimulation intensity was increased until ≈10% of the responses recorded were suprathreshold. Following IGF-I (Preprotech) was added to the bath and the PSPs were recording for 15 min. Then, we switched back to voltage-clamp, returning the intensity and frequency to that used for the baseline recording, and we recorded the PSCs for 20 min. After that, we washed the IGF-I and recorded PSCs during 20 min after washout IGF-I. We switched each 10 min from voltage-clamp to current-clamp to check the amplitude of the PSPs (during 2 min) after IGF-I washout. We added different IGF-I concentration (5, 7 and 10 nM) Plots of the changes induced by IGF-I on EPSCs peak amplitude (percentage from control) versus time were constructed (Figure 2). The same experiment was performed in present to the IGF-I antagonist receptor 7-[cis-3-(1-178azetidinylmethyl)cyclobutyl]-5-[(3-phenylmethoxy)phenyl]-7H-pyrrolo[2,3-]pyrimidin-179 4-amine (NVP-AEW 541, 40 μM; Cayman Chemicals) plus IGF-I. In some experiments, the intracellular solutions could also either contain: 1,2-Bis(2-aminophenoxy) ethane-N,N,N′,N′-tetra acetic acid (BAPTA; 20 mM) or light chain of the B-type botulinum toxin (BOTOX, 1 μM) (Figure 3).

Synaptic stimulation increased (SSI), after 5 min of stable baseline of EPSCs in voltage-clamp, the recording was switched to the current-clamp and the stimulation intensity was increased to values in which an EPSP followed by an AP was triggered during 15 min at 0.2 Hz. Next, the values of synaptic stimulation were returned to control conditions and PSCs were recorded for 25 min (Noriega-Prieto et al., 2019) (Figure 4).

Simulated spike, after 5 min of stable baseline of EPSCs in voltage-clamp, after that, recording mode was switched to the current-clamp and a previously obtained neuronal spike was used as stimulus to depolarize the neurons through the recording pipette during 15 min at 0.2 Hz. Next, we were returned to control conditions and EPSCs were recorded for 25 min.

After deep anesthesia with sodium pentobarbital (60 mg/kg), 6 month-old male mice (C57BL6/J) were perfused transcardially with 0.1 M phosphate-buffered saline (PBS), pH 7.4 followed by 4% paraformaldehyde, 75 mM lysine, 10 mM sodium metaperiodate in 0.1 M PB, pH 7.4. Brains were removed, post-fixed overnight in the same fixative solution at 4°C, coronally sectioned at 50 μm thicknesses on a vibratome (Leica VT1000S), and serially collected in wells containing cold PB and 0.02% sodium azide.

For IGF-IR immunogold labeling, sections containing the somatosensory cortex were used. Sections were first washed with PBS and incubated in a 50 mM glycine solution 5 min to increase antibody binding efficiency. Following a standard immunocytochemical protocol, tissue was first free-floating incubated in a rabbit polyclonal anti-IGF-IRα antibody (1/250, Santa Cruz) in a PBS 0.1 M/1% BSA solution for 48 h at 22°C. Then, sections were washed in PBS, and incubated with 1.4 nm gold-conjugated goat anti-rabbit IgG (1:100; Nanoprobes) overnight at 22°C. After post-fixing with 2% glutaraldehyde and washing with 50 mM sodium citrate, labeling was enhanced with the HQ Silver^™ Kit (Nanoprobes), and gold toned. Finally, immunolabeled sections were fixed in 1% osmium tetroxide, block stained with uranyl acetate, dehydrated in acetone, and flat embedded in Araldite 502 (EMS, United States). Selected areas were cut in ultrathin sections (70–80 nm) and examined and photographed with a JEOL JEM1400 electron microscope. As a control for the immunogold technique, sections were processed as above but omitting the primary antibody. No specific labeling was observed in these control sections. We identified excitatory and inhibitory synapses based on the morphological appearance of the PSD in EM images. Excitatory (asymmetric) synapses typically display a pronounced post-synaptic density (PSD) much thicker than the relatively faint presynaptic thickening, while the PSD in inhibitory (symmetric) synapses looks similar to the presynaptic membrane (any synapse with a less marked PSD, similar to the presynaptic thickening, was classified as “symmetric”) (Gray, 1969).

C57BL6/J mice (15 days old) were anaesthetized with pentobarbital (60 mg/kg) and perfused transcardially with 4% paraformaldehyde in 0.1 M phosphate buffer pH 7.4 (PB). The brain was then removed and post-fixed overnight. Coronal 50 μm-thick sections were cut and collected in PBS. Free-floating brain sections were pre-incubated in PB with Triton X-100 1% and bovine serum albumin 1%, and incubated overnight at 4°C with the respective primary antibody [mouse anti-PSD95 (1:500), mouse anti-vGAT (1:50) and rabbit anti-IGF-IR (1:100)]. Incubations were performed in two steps (first, PSD95/vGAT and then IGF1R). The following primary antibodies were utilized: a mouse anti-PSD95 antibody (Thermo Scientific 1:500), a mouse vGAT (Synaptic System 1:50) and a rabbit anti-IGF1R antibody (Santa Cruz, 1:100). Secondary Alexa-labeled antibodies from Molecular Probes (Donkey anti-mouse 488 Alexa-conjugated for the PSD95 and vGAT detection and donkey anti-mouse 594 Alexa-conjugated for the IGF1R detection) were used at a final concentration of 1:1000. All sections were counterstained with Hoechst (Calbiochem, 1:500). The incubation periods used were 1 h at room temperature (RT) and 48 h at 4°C for primary antibodies, 1 h RT and 24 h at 4°C for secondary antibodies, and 15 min for Hoechst incubation.

Pre- or post- synaptic origin of the synaptic plasticity induced by IGF-I was tested by analyzing the modification in the variance that parallel the synaptic current amplitude change, which reflect the change in transmitter release probability (Clements, 1990; Malinow et al., 1990; Kuhnt and Voronin, 1994). To estimate the modification of the synaptic current variance, we first calculated the noise-free coefficient of variation (CV_NF) of the synaptic responses before (control conditions) and during IGF-I. We used the formula CV_NF = √ (δ_{synaptic current}² − δ_noise²)/m, where δ_noise² and δ_{synaptic current}² are the baseline and synaptic current peak variance, respectively, and m is the mean peak amplitude of the synaptic current. The ratio of the CV_NF (CV_R) before (control conditions) and during IGF-I was obtained then for each neuron as CV_IGF-I/CV_control (Clements, 1990). We constructed plots comparing variation in the normalized m (termed M) to the change in 1/CV_R² in each cell (Malinow et al., 1990). In these plots, depression of the synaptic currents has a presynaptic origin when values are below or in the diagonal, whereas points above the diagonal indicate a postsynaptic origin. However, potentiation of synaptic currents has a presynaptic origin when values are above or in the diagonal, whereas points below the diagonal indicate a postsynaptic origin (Faber and Kornt, 1991). This method requires a binomial EPSC amplitude distribution, a condition that must be met for the synaptic variance to reflect the probability of transmitter release. We could not directly test whether our data fitted a binomial distribution, but synaptic fluctuations were always evident and we assumed that synaptic release followed a binomial distribution. Data analysis was done in Clampfit 10 (Axon Instrument) and graphs were drawn in SigmaPlot 11. In all cases, statistical estimates were made with student’s two-tailed paired t-tests, and data are presented as means ± SE. To analyze the effects of different treatments and conditions, we carried out multiple comparison testing between the different groups using one-way ANOVA. The post hoc test used was Holm–Sidak or Dunn’s for parametric or non-parametric values respectively, versus control comparisons using the “after stimulus” situation and the “basal” condition as the controls to compare with a Holm–Sidak’s or Dunn’s multiple comparisons test. Statistical differences were established with p < 0.05 (∗), p < 0.01 (∗∗) and p < 0.001 (∗∗∗) for student’s t-test and p < 0.05 (#), p < 0.01 (##) and p < 0.001 (###) for the post hoc Holm–Sidak and Dunn’s test. The sample size is shown as n = x/y (x shows the number of cells, and y shows the number of animals).

We have previously shown that 10 nM IGF-I decreases the induction threshold of LTP_H (Noriega-Prieto et al., 2021). Here, we analyzed the effect of a slightly lower dose of IGF-I on LTP_H induced by spike timing dependent plasticity (STDP) at layers 2/3 of the barrel cortex (Figure 3A). We first checked in control conditions, the STDP protocol, consisting in a subthreshold PSP followed by a back-propagating action potential (BAP) at delays of 10 ms repeated 50 times at 0.2 Hz (Figure 3B) and then we performed similar experiments in slices in which previously IGF-I at doses of 10 nM, 7 nM or 5 nM was bath applied during synaptic stimulation of layer 4 at 0.2 Hz (Figure 3C). In control conditions, the STDP protocol induced an LTP_H of PSPs (from 96.01 ± 4.12 to 137.62 ± 8.33% of amplitude, p < 0.001; n = 7/3; Figures 3D,G, LTP control). However, in slices pretreated with 10 nM IGF-I, the STDP protocol induced a more robust LTP_H of PSPs (from 99.36 ± 1.08 to 141.59 ± 4.38% of amplitude after 20 pairings, p < 0.001; n = 6/2 and from 99.36 ± 0.89 to 169.13 ± 6.36% of amplitude after 50 pairings, p < 0.001; n = 6/3; Figures 3E,G, LTP facilitated). Conversely, in slices pretreated with 7 nM IGF-I, the STDP protocol did not induce LTP_H of the PSPs (from 101.66 ± 1.12 to 99.85 ± 6.35% of amplitude after 50 pairings, p > 0.05; n = 6/3; Figures 3F,G, LTP impairment). Interestingly, 7 nM IGF-I with 100 pairings repetition induced an LTD (from 102.74 ± 1.26 to 72.65 ± 5.19% of amplitude after 100 pairings, p < 0.001; n = 5/2; Figures 3F,G, LTP impairment), which further supports the idea of different IGF-I levels differently regulate the synaptic plasticity. Additionally, 50 pairings of pre- postsynaptic neuronal activity in the presence of picrotoxin (50 μM) and CGP (1 μM) to isolate the excitatory synaptic transmission, after 10 nM IGF-I did not induce any modulatory effect in the excitatory postsynaptic currents (EPSCs) (from 100. 57 ± 0.63 to 105.25 ± 10.13% of amplitude after 50 pairings; p > 0.05, n = 6/3; Supplementary Figures S2A–D, 50 pairings). Thus, only 10 nM, but not 7 nM IGF-I favored the induction of LTP_H through the modulation of inhibitory synaptic transmission (Noriega-Prieto et al., 2021). In other words, the activation of IGF-IRs favors or impairs LTP_H depending on IGF-I concentration.

We next compared the effects of these two doses of IGF-I on synaptic transmission. For this set of experiments, we recorded at L2/3 PNs (Figure 1A) the postsynaptic potentials (PSPs, Figure 1B) and currents (PSCs, Figure 1D top) evoked by stimulation of layer 4. After 5 min of recording PSPs and PSCs, 10 nM or 7 nM IGF-I was applied during 35 min. As previously published (Noriega-Prieto et al., 2021), 10 nM IGF-I induced a long-term potentiation (LTP) of the PSPs (termed LTP_IGF-I) that remained 30 min after IGF-I washout (from 100.70 ± 1.58 to 127.50 ± 8.06% of amplitude at 60 min after IGF-I, p < 0.01, n = 8/3; Figures 1B,C, 10 nM IGF-I). However, the PSCs were transiently potentiated, returning to control values after IGF-I washout (from 98.58 ± 0.52 to 100. 29 ± 5.89% of amplitude 60 min after IGF-I, p > 0.05, n = 9/3; Figure 1D, IGF-I 10 nM, white circles). Interestingly, 7 nM IGF-I induced a long- term depression (LTD) of the PSPs (from 104.46 ± 0.82 to 70.10 ± 5.20% of amplitude 60 min after IGF-I, p < 0.01, n = 5/2; Figures 1B,C, 7 nM IGF-I) and the EPSCs (termed LTD_IGF-I from 103.78 ± 1.23 to 66. 22 ± 9.21% of amplitude 60 min after IGF-I, p < 0.05, n = 5/2; Figure 1D black circles and 7 nM IGF-I). Moreover, neither the PSPs (from 99.37 ± 0.67 to 102. 74 ± 2.31% of amplitude 60 min after IGF-I, p > 0.05, n = 5/2; Figures 1B,C, 5 nM IGF-I) nor PSCs (from 101.17 ± 0.88 to 91.95 ± 4.60% of amplitude 60 min after IGF-I, p > 0.05, n = 6/3; Figure 1D grey circles, and IGF-I 5 nM) were modified when 5 nM IGF-I was bath perfused. These results demonstrate that IGF-I can induce LTP or LTD of the PSPs at L2/3 PNs of the barrel cortex in a concentration dependent manner.

We next analyzed the locus of expression of IGF-I mediated LTD of the EPSCs. We first studied whether it was paralleled by a decrease in the probability of glutamate release. We pharmacologically isolated the EPSCs by blocking GABA_A inhibition with PiTX, and applied 7 nM IGF-I (Supplementary Figure S1). Under these conditions, IGF-I induced a similar LTD of the ESPCs than IGF-I 7 nM, that lasted at least 40 min of recording (from 99.59 ± 0.44 to 67.36 ± 3.87% of amplitude, p < 0.001, n = 6/2; Supplementary Figure S1A, black circles). This effect was inhibited with the IGF-IR antagonist NPV-AEW 554 (from 100.37 ± 0.37 to 94.87 ± 3.67% of amplitude, p > 0.05, n = 5/2; Supplementary Figure S1A, white circles). To investigate the pre or postsynaptic locus of expression of this LTD, we analyzed the effect of IGF-I on the pair pulse ratio (PPR) and on the EPCS variance by measuring the coefficient of variation (CV). The effect of IGF-I on both the PPR (from 0.98 ± 0.01 to 1.28 ± 0.10 before and after IGF-I respectively, p < 0.05, n = 6/2; Supplementary Figure S1B left) and the analysis of 1/CV² plots (linear correlation R² = 0.974, Supplementary Figure S1B right) revealed that LTD_IGF-I was due to a decrease in the probability of release of glutamate.

Neurons secrete IGF-I by an activity-dependent pathway of exocytosis, and a mild depolarization is sufficient to induce IGF-I secretion in olfactory bulb neurons (Cao et al., 2011). Therefore, we next tested whether changes in cytosolic Ca²⁺ levels of postsynaptic PNs were involved in IGF-I potentiation of the EPSC. We carried out similar experiments as before, but in the presence of the Ca²⁺ chelator BAPTA (20 mM) in the recording pipette (Figure 2A). Under these conditions, 10 nM IGF-I induced an EPSCs depression (43.90 ± 4.04% of baseline, p < 0.001, n = 6/3, Figure 2B, 10 nM IGF-I, green circles) rather than an EPSCs potentiation observed in the absence of BAPTA. This EPSC depression was absent with 7 nM IGF-I (from 98.79 ± 1.37 to 99.19 ± 2.23% of amplitude, p > 0.05, n = 6/3; Figure 2B, 7 nM IGF-I, blue circles). Moreover, the plasticity was abolished in the presence of the IGF-I receptor antagonist NPV-AEW 554 (from 99.42 ± 1.79 to 104.08 ± 4.39% of amplitude, p > 0.05, n = 5/2; Figure 2B, 10 nM IGF-I + NVP, light red circles). These results indicate that IGF-I mediate EPSCs potentiation or depression depending on the cytosolic Ca²⁺ level, being all these forms of synaptic plasticity dependent on the activation of IGF-IRs.

Synaptotagmin 10 (Syt10) acts as the Ca²⁺-sensor that triggers IGF-I exocytosis in olfactory bulb neurons (Cao et al., 2011). Thus, we prevented exocytosis by using the light chain of the B type botulinum toxin (i.e., Botox 0.5 μM), which inhibits the SNARE protein-mediated membrane fusion of endosome complexes, and tested whether IGF-I effects depend on exocytosis. Surprisingly, 7 nM IGF-I did not modulate the EPSCs (data not shown), while IGF-I 10 nM depressed the EPSCs under BOTOX (42.26 ± 8.63% of baseline, p < 0.01, n = 6/3; Figure 2C, BOTOX + IGF-I 10 nM), suggesting that higher IGF-I concentrations are required for LTD_IGF1 under Botox. Interestingly, increasing IGF-I to 20 nM was able to induce the potentiation of the EPSCs (227.80 ± 20.43% of baseline, p < 0.001, n = 7/3; Figure 2C, BOTOX + IGF-I 20 nM) indicating that higher IGF-I concentrations are required to LTP_IGF1 under Botox probably because it blocked the activity-dependent release of IGF-I from the postsynaptic neuron.

Since cytosolic calcium levels and exocytosis are determinant in the induction of LTD and LTP by IGF-I, we next tested whether increases of cytosolic calcium induced by synaptic stimulation could be enough to induce IGF-I-mediated synaptic plasticity. After 5 min of recording the EPSCs, we increased the intensity of synaptic stimulation (SSI, synaptic stimulation increase) until a PSP followed by an action potential was recorded (Figures 4A,B). Next, we maintained evoked these suprathreshold responses for 15 min by SSI, and then we turned the stimulation intensity back to control values. This protocol of stimulation induced a LTD of the EPSCs (from 101.35 ± 2.43 to 54.54 ± 8.43% of amplitude, p < 0.01, n = 5/2; Figures 4A,B, black circles, SSI) that was prevented with NVP (from 101.09 ± 0.91 to 94.33 ± 4.50% of amplitude, p > 0.05, n = 6/3; Figures 4A,B, white circles, SSI + NVP), or under Botox (from 100.33 ± 1.01 to 100.60 ± 2.43% of amplitude, p > 0.05, n = 6/3; Figures 4A,B, red circles, SSI + Botox). These results suggest that suprathreshold responses induced by SSI are sufficient to induce an IGF-I-mediated LTD of the EPSCs dependent on endosome exocytosis. Finally, we checked whether simulation of these suprathreshold responses (Figures 4C,D, Simulated Spike) by injecting them through the patch recording electrode could induce a similar LTD of the EPSCs. As shown in Figures 4C,D, simulation of suprathreshold responses for 15 min was enough to induce a presynaptic LTD of the EPSCs (40.56 ± 3.58% of baseline, p < 0.01, n = 5/2; Figures 4C–E purple circles, simulat. spike), that is prevented with NVP (3.25 ± 3.38% of baseline, p > 0.05, n = 5/2; Figures 4C,D blue circles, simulat. spike + NVP). These results suggest that suprathreshold responses can induce increases in postsynaptic calcium levels that trigger the release of IGF-I and subsequent induction of LTD of the EPSCs.

We finally analyzed whether IGF-IRs were present at presynaptic terminals of excitatory in the postsynaptic L2/3 PNs. Electron micrographs demonstrated IGF-IR silver-enhanced immunogold labeling (Figures 5A,B, white arrows) at presynaptic terminals of both excitatory (Figure 5A) and inhibitory (Figure 5B) synapses. Moreover, Double immunocytochemical analysis for pre (vGAT)- and post- (PSD95) synaptic markers and IGF-IR indicate the presence of immunopositive puncta opposing each other in close proximity (Figures 5C,D), in agreement with the presence of IGF-IR in both pre- and post-synaptic terminals. The presence of IGF-IR in the presynaptic terminals of both glutamatergic and GABAergic synapses supports the presynaptic LTD of the EPSCs induced by 7 nM IGF-I (present results) and the presynaptic LTD of the IPSCs generated by 10 nM IGF-I (Noriega-Prieto et al., 2021).

In the central nervous system, IGF-I/IGF-IR signaling is critical for experience-dependent synaptic and neuronal plasticity in sensory cortices, adult neurogenesis (LLorens-Martín et al., 2010), synaptic vesicle release, and neuronal excitability (Nuñez et al., 2003; Maglio et al., 2021). Our present results challenge the standard view that IGF-I favors cognition by inducing LTP in cortical circuits, and it expands the range of brain IGF-I actions. Thus, we provide novel evidence that IGF-I levels determine the sign of plasticity induced at the barrel cortex; high levels induce LTP, while lower levels induce LTD, which in turn, favor or impair Hebbian LTP, respectively (Figure 6). While the reduction in the inhibitory tone by 10 nM IGF-I and the subsequent LTP of the synaptic plasticity reduces the threshold for inducing Hebbian LTP, the reduction of the excitatory transmission induced by 7 nM IGF1 would increase the threshold maybe by an insufficient activation of the necessary number of NMDARs to increase the cytosolic calcium to induce synaptic plasticity. Although the observation that IGF-I action on the brain is crucial for learning and memory is well established, here we are proposing an additional interpretation of the classical IGF-I mechanism. These results indicate that brain levels of IGF-I play an important role in synaptic plasticity, which is crucial for memory processes. Therefore, any changes in brain IGF-I levels could impact synaptic plasticity and cognitive functions.

Extracellular IGF-I activates IGF-IRs under resting conditions, maintaining basal transmission and ongoing spiking activity in the physiological range, and induces synaptic modulation (Gazit et al., 2016). Presynaptic IGF-IRs are basally active, thus regulating glutamatergic synaptic transmission by modulating the glutamate release probability (Gazit et al., 2016). In fact, it can be concluded that tonic release of IGF-I and subsequent activation of IGF-IRs modulates synaptic vesicle release, leading to a short term depression in excitatory hippocampal neurons (Gazit et al., 2016). In agreement with these results, we also observed a long-term depression of synaptic transmission in the absence of postsynaptic activity (when firing at the recorded PN was prevented in voltage clamp) or when BAPTA abolished the cytosolic calcium increase mediated by this activity at PNs. Under Botox 7 nM IGF-I is not sufficient to induce LTD and 10 nM IGF-I is required for the LTD induction, whereas increasing the concentration to 20 nM IGF-I induces LTP. This demonstrates that endosome fusion is critical for establishing the concentration threshold for the induction of LTD and LTP by IGF-I, and suggests that exogenous application of IGF-I could induce neuronal-released IGF-I stored in endosomes (a calcium and endosome dependent IGF-I induce IGF-I release). These results strongly indicate that the tonic modulation of synaptic transmission induced by IGF-I described at the hippocampus is also present at glutamatergic synapses of the barrel cortex.

Our previous results show that 10 nM IGF-I induces an LTP of the PSPs, that we have termed LTP_IGFI (Noriega-Prieto et al., 2021). Moreover, we demonstrated that IGF-IR activation favors the induction of NMDAR-dependent LTP and improves texture discrimination of the mouse whiskers. Here we show that lower IGF-I levels (7 nM) induce LTD of the PSPs, that we have termed LTD_IGFI and impairs the induction of NMDAR-dependent LTP. These results are supported by the presence of IGF-IR in the presynaptic terminal of both excitatory and inhibitory synapses described in this work. In addition, based on the evidence, this mechanism of action has been studied in the hippocampus, where IGF-I can induce the release of GABA to regulate endogenous ACh release, possibly acting via GABAergic neurons (Seto et al., 2002). Moreover, olfactory learning in a social context selectively induces LTP of the GABAergic component of reciprocal synapses between granule and mitral cells in the medial olfactory bulb (MOB), requiring an autocrine and/or paracrine action of IGF-I to enhance postsynaptic GABA receptor function. Indeed, blocking Ca²⁺-triggered IGF-I release prevents GABAergic LTP (Liu et al., 2017). At any rate, our observations contribute to the notion that IGF-I modulates both excitatory and inhibitory synaptic activity throughout the CNS (Maglio et al., 2021; Noriega-Prieto et al., 2021). According to our results, IGF-I is able to induce a dual effect on glutamatergic synaptic transmission by the activation of IGF-IRs. In fact, both a pre-synaptically IGF-I-mediated potentiation and depression of the EPSCs were observed, and both were prevented by NPV-AEW 554 (Maglio et al., 2021; Noriega-Prieto et al., 2021).

On the other hand, we also observed that IGF-I induced an increase in glutamatergic synaptic transmission through the activation of IGF-IRs. As discussed above, IGF-I-induced potentiation of excitatory synaptic transmission is dependent on the presence of exogenously applied IGF-I, thereby pointing to the importance of reaching specific local levels of IGF-I. Indeed, only IGF-I mediated depression of the EPSCs are observed by the release of IGF-I induced by high synaptic stimulation or simulated spikes (Figure 4). These results points to the importance of IGF-I uptake from the plasma in the induction of bidirectional plasticity, since only depression of the synaptic transmission can be induced without extracellular perfusion of IGF-I. Whereas there is a tonic IGF-I release that induces depression of glutamatergic synaptic transmission (Gazit et al., 2016), our results suggest that the bidirectional effect of IGF-I on the modulation of the EPSCs may depend on the levels of IGF-I reached (Figure 6). An IGF-I -mediated EPSC depression is produced when PNs fires during supra-threshold responses, whereas an IGF-I-mediated EPSC potentiation is induced by bath applied IGF-I. Under physiological conditions, higher levels of IGF-I could be reached in a neuronal-activity dependent manner (Nishijima et al., 2010). In fact, we described that serum IGF-I input to the brain is regulated by an activity-driven process that includes increased blood-brain barrier permeability to serum IGF-I (Nishijima et al., 2010). Also, it has been demonstrated that physical exercise induces increased brain uptake of serum IGF-I by specific groups of neurons throughout the brain (Carro et al., 2000; Nuñez et al., 2003). This increase in the uptake of IGF-I, under both mentioned physiological circumstances, induces an increase in neuronal excitability, which perfectly correlates with our previous in vivo results (Noriega-Prieto et al., 2021). The ex vivo investigation showed here suggests that this increase in IGF-I uptake would be essential in EPSC potentiation by increasing local IGF-I levels to those required for the modulation of synaptic transmission, playing a role in the increase in cortical activity.

In summary, IGF-I induces bimodal regulation of the excitatory and inhibitory synaptic transmission depending on its levels. This bidirectional action probably contributes to favor or impair the generation of associative memories impacting the barrel cortex-related behaviors. Although there is a SNARE dependent release of IGF-I from neurons (Cao et al., 2011) and Botox inhibits the SNARE protein-mediated membrane fusion of endosome complexes, the application of Botox, though effective in blocking the release of IGF-I stored in endosomes, can affect other neuromodulators stored in endosomes that modulate synaptic transmission, such as BDNF (Nasrallah et al., 2021), and interfere with glutamate receptor trafficking (Pellett et al., 2015). Additionally, further studies are necessary to investigate whether the bidirectional modulation induced by IGF-I is maintained in older animals. Despite potential limitations, our previous findings, together with the present work, reveal novel insights into the mechanisms of IGF-I signaling in the cortex.